Is the punctate labelling in the images your protein of interest with the fuzzy labelling what you think is the background labelling? Or is everything fluorescing in the two images what you think is background?

Have you tried dilution series of primaries and secondaries? Reducing the gain/laser power on the microscope?

The other secondaries you tried, were they different fluorophores or different hosts but the same fluorophore?

Out of interest, are you permeabilising the sections prior to labelling? You sometimes get differential labelling depending on your permeabilisation technique - eg triton X100 vs saponin.

It could just be the unfortunate case of your rabbit primaries cross-reacting with your samples... have you tried contacting the supplier for help or are these primaries that you've raised yourself?

Hi John,

The larger oval shapes are the protein of interest, which is a transcription factor and hence labels the nuclei of certain neurons here. My problem is the ca 1-pixel large dots (much smaller than the ovals) that are spread everywhere. These dots are not so obvious on these images actually, but is much worse when viewed in the eyepiece, or when taking a confocal image. Reducing primary concentration reduces the signal of the protein of interest. I could try to reduce secondary concentration, but suspect the protein of interest signal would also suffer. I tried reducing laser gain, but these dots are some of the most fluorescent signal. I tried using secondaries from both goat or donkey, and tried alexa 555, alexa 488 and cy3 from invitrogen and jackson suppliers. I permeabalize with TBS + 0.1 % tween 20. All rabbit primaries that I've had this problem with are from commercial sources. I have not tried asking them about it.

I just tried some new experiments, and Im still completely puzzeled by the issue. I tried blocking with 4% BSA in TBST, which I think reduced the dots a lot, but the blocking was insufficient, and my real signal was no longer strong. However, my controls without primary gave no background or dot signal with either 4% BSA or 10% donkey serum blocking. So I cant understand what else it can be but some kind of reaction between the primary and the blocking, or the primary and the mouse brainstem tissue. But then it is very strange that this occurs with all different rabbit-primaries I try, and not with other primaries. Maybe it just so happens that most of my rabbit primaries are bad/dirty? I also tried preincubating secondary antibodies with aceton-powdered mouse-brainstem, but dots were still seen after that. 

Dear Anders,

You have done all that I think you can to trouble-shoot whether it is a problem with the secondary, and I agree with you that it must be something about your rabbit primary antibodies (what a nightmare!).  Do you also have proteins to block non-specific binding of the primary antibody in the diluent that you use for labeling?

Playing around with the blocking protocols or diluents might help, since changing the blocking reagent does seem to be reducing some of the punctate staining. I have included links here to two of my publications with different blocking protocols that were provided by my collaborators to insure clean labeling using their polyclonal antibodies.  If you decide you want to try them, I suggest using the recipes in the anti-CNTF-R alpha paper first, since that was clean even with the use of FITC tyramide amplification of ABC elite reagent.  The recipes in the anti-GAP43 paper are much more involved, but they do eliminate background labeling very nicely.  I sometimes use those recipes to get clean background for tract tracing in goldfish.

If you need the protocols, rather than just the papers, please let me know.

Good Luck!

Jill

CNTF-R alpha: http://onlinelibrary.wiley.com/doi/10.1002/cne.21174/full

Gap-43: http://www.sciencedirect.com/science/article/pii/0306452294905525

Hi Jill,

Thank you for your comment! Yes, I use the same buffer for blocking and for primary incubation, usually 10% donkey serum. 

I checked out the papers you linked to, with regards to blocking. The CNTF-R paper appeared to involve much of the same steps as I'm already doing. The changes were using tritonX 100 instead of tween 20, and at a slightly higher concentrations, more washing steps, and also a small amount of BSA in the blocking buffer. Do you think these changes might be enough to remove or reduce my unspesific staining? For the Gap-43 paper I could not see any description of the blocking procedure. Could you point me in the right direction? Or better yet, could you send me the protocol (lunde.anders AT gmail.com)? I am very interested in the protocol since you said it eliminates backgorund labeling very nicely! By the way, are you then also referring to grainy/dotty bakground staining like I'm seeing?

Thanks in advance!

Hi Hussein,

Thank you fo providing your protocol. I would be interesting to try permeabilizing with 1% triton, and using only 1% serum in the blocking buffer. Is 1% usually sufficiant to block unspecific staining? 

Is 0.1 M PBS used in all steps described? Why and what is the difference between using 0.1 and 1 M PBS? 

Best,

Anders

You should not use short fixation times (if the 30 min is time of fix). 4% paraformaldehyde is a very very slow fixative.  Use of longer fixation times could help.  Most likely the problem is that you do not have the optimal antibody staining conditions and use of fluorescent-labeled secondaries is the least sensitive of any method available. For embryonic tissue you are dealing with tissue that is mostly water. This too can be why better fixation is needed. If you were to amply your signals using less antibody, you could get cleaner staining. In my experience, addition of normal serum does nothing to help background.

Hi. Just some update from me. I have not gotten to try any of your various protocol advice yet.

I'm running a parallel discussion here: http://www.protocol-online.org/forums/topic/35658-getting-terrible-grainy-background-in-immunofluorescence-experiment/

And I've gotten an advice about spinning primary solution. I usually do this, but I think I often go to the bottom of the aliquot to pipet out after this, due to small aliquot sizes. Maybe this is an important factor, I need to avoid.

And some clarifications: 

I always use the same imaging/visualizing setup for "red" channel, across different antibodies. For example I used a goat-antibody1 and a rabbit-antibody2 in a parallell experiments, label both with Cy3-secondaries, and the goat one is beautiful with 0 background, while the rabbit is really bad. And like I said, the bad rabbit staining happens with 4 different secondaries I've tried. The problem is also very apparent in at least 3 of 4 rabbit primaries I have, but almost never seen in 10-20 other primaries I use from different species.

After reviewing all my tests and controls it is clear to me that this phenomena is sensitive to tissue batches (I've run different tissue batches in prallel, using otherwise same conditions, with a really marked difference). I try to treat each tissue preparation similar, so it must be rather sensitive. FYI it is embryonic mouse or chicken brainstem preparations that are labeled with fluorescine-dextran and incubated in ACSF for ~9 hours, before 30 min 4% PFA fix in cold, 20-30% sucrose treated, OCT embedded and cryosectioning. Storage if OCT blocks, cryosections, ACSF composition are possible important parameters. 

I was a bit unclear before, but multiple tests show that my controls which exclude primary antibody never gives any background. 

My only hope now is to make a few new batches of tissues, and hope they will be better. Also possibly take more care with the spinning of primaries.

I have the same problem, but for me it's mostly in green. Any suggestions for the final solution?

Cagla, I doubt that it's a novel localization thing, since the grainy spots don't colocalize with DAPI or anything else. At least, that's the case for me. @cagla akay

No I never fixed it, just went with it. Its good enough.

We have trouble with green autofluorescence and yesterday for the firs time tried Vector's Trueview autnofluorecence quencching kit and it worked like a charm. Definitely look into it. It's pretty cheap. The first image is with nothing added and the second is with the quencher. They were taken at the same exposure but the one with the quencher is totally black!!

You are likely using the wrong concentration of antibodies

use Tissufix solution - they say it is perfect designed in such a way that the alcolic solution is balanced with the polymerization solution.

and Sodium borohydride for reduction of background autofluorescence in sections.

Yes I would suggest autofluorescence

could also be the concentration of primary antibody is too high, you could also try a low incubation temp eg 4C

Hello Anders Lunde were you able to figure it out? I am having similar issue with my rabbit antibodies